Optimizing DNA double strand break repair for homologous recombination based gene therapy

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Optimizing DNA double strand break repair for

homologous recombination based gene therapy

zur Erlangung

des Akademischen Grades

Doktor der Naturwissenschaften

Dr. rer. Nat.

Im Fachbereich Biologie und Chemie

der Justus-Liebig-Universität Gießen

vorgelegt von

Song, Fei

aus Shanghai

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Die vorliegende Arbeit wurde in der Arbeitsgruppe Experimentale Ophthalmologie

der Klinik und Poliklinik für Augenheilkunde des Fachbereichs 11 der

Justus-Liebig-Universität Gießen, in der Zeit von November 2013 bis November 2016,

unter der Leitung von Prof. Dr. Dr. Knut Stieger angefertigt.

Erstgutachter: Prof. Dr. Peter Friedhoff

Institute of Biochemistry

Heinrich-Buff-Ring 17

35392 Gießen

Zweitgutachter: Prof. Dr. Dr. Knut Stieger

Department of Ophthalmology

Friedrichstr.18

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Erklärung

Hiermit versichere ich, die vorliegende Dissertation selbständig und ohne fremde

Hilfe verfasst zu haben und keine anderen als die hier angegebenen Hilfsmittel

benutzt zu haben. Alle Textstellen, die wörtlich oder sinngemäß aus

veröffentlichten Schriften entnommen sind, und alle Angaben, die auf mündlichen

Auskünften beruhen, sind als solche kenntlich gemacht.

Gießen,

_____________

(Fei Song)

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Publications

Slijkerman, R. W.*, F. Song*, G. D. Astuti*, M. A. Huynen, E. van Wijk, K. Stiegerand R. W. Collin (2015). "The pros and cons of vertebrate animal models for functional and therapeutic research on inherited retinal dystrophies." Prog Retin Eye Res 48: 137-159.

*Three first authors have contributed equally to this work.

Yanik, M., B. Muller, F. Song, J. Gall, F. Wagner, W. Wende, B. Lorenz and K. Stieger (2016). "In vivo genome editing as a potential treatment strategy for inherited retinal dystrophies." Prog Retin Eye Res. (In press)

F. Song, K. Stieger (2016). “Optimizing the DNA donor template for homology directed repair

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Abbreviations

% percent °C celsius µ micro µl microlitre 3’ three prime 5’ five prime 53BP1 p53 binding protein 1 A adenine aa amino acid

AAP assembly-activating protein

AAV adeno-associated virus

ADA-SCID adenosine deaminase deficiency-severe combined immunodeficiency

AP alkaline phosphatase

ATM ataxia-telangiectasia mutated

ATR ataxia-telangiectasia RAD3-related kinase

BFP blue fluorescent protein

BIR break-induced replication

bp base pair

BRCA1 breast cancer type 1

BRCA2 breast cancer type 2

BTR BLM-TOPOIIIα-RMI1-RMI2

C cytosine

Cas CRISPR-associated

CDK cyclin-dependent kinase

cDNA complementary DNA

c-NHEJ classical nonhomologous end joining

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Abbreviatations

VI

CRISPR clustered regularly interspaced short palindromic repeats

crRNA CRISPR RNA

CtIP CtBP-interacting protein

ddH2O double-distilled water

DDR DNA damage response

del deletion

dH2O distilled water

DMEM Dulbecco's Modified Eagle Medium

DMSO dimethyl sulfoxide

DNA deoxyribonucleic acid

DNA-PK DNA-dependent protein kinase DNA-PKcs DNA protein kinase catalytic subunit

dNTP deoxynucleotidetriphosphates

DMD duchenne muscular dystrophy

DSB double-strand break

DSBR double-strand break repair

Fig figure

G guanine

GFP green fluorescent protein

gRNA guide RNA

HDR homology-directed repair

HNSCC head and neck squamous cell carcinoma

HR homologous recombination

IDLV integration-deficient lentiviral vector indel insertion and deletion

IRD inherited retinal dystrophy

kb kilo base

lin. linearized

L liter

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Abbreviatations

VII

LCA Leber's congenital amaurosis

LOH loss-of heterozygosity

LPL lipoprotein lipase

LPLD lipoprotein lipase deficiency

LV lentivirus

MDC1 mediator of DNA damage checkpoint 1

min minute

ml milliliter

MMEJ microhomology-mediated end-joining

MRN Mre11-Rad50-Nbs1

mRNA messenger RNA

mut mutated

NGS next generation sequencing

NHEJ nonhomologous end-joining

NLS nuclear localization signal

Nr. Number

OPEN Oligomerized Pool ENgineering

pla plasmid

PAM protospacer adjacent motif

PBS phosphate-buffered saline

PCR polymerase chain reaction

polθ DNA polymerase theta

rAAV recombinant adeno-associated virus RAP80 receptor-associated protein 80

RFLP restriction-fragment length polymorphism

RFP red fluorescent protein

RFP168 ring finger protein 168

RFP8 ring finger protein 8

RGN RNA-guided endonuclease

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Abbreviatations

VIII

RPA replication protein A

RP retinitis pigmentosa

RPGR retinitis pigmentosa GTPase regulator

RT room temperature

RVD variable di-residue

SDSA synthesis-dependent strand annealing

SIN self-inactivating

siRNA small silencing RNA

SOC Super Optimal broth with Catabolite repression SpCas9 Streptococcus pyogene Cas9

SSA single-strand annealing

SSB single-strand break

ssDNA single-stranded DNA

T thymine

TALE transcription activator-like effector

TALEN transcription activator-like effector nuclease

TBE Tris-Boric acid-EDTA

TLR traffic light reporter

TLS translesion synthesis

tracrRNA transactivating CRISPR RNA VEGF vascular endothelial growth factor XLRP X-linked retinitis pigmentosa

XRCC4 X-ray repair cross-complementing protein 4

ZF zinc-finger

ZFN zinc-finger nuclease

α alpha

β beta

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List of figures

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List of figures

Fig. 1: Generation of recombinant AAV vectors………..5

Fig. 2: Recombinant lentivirus vectors………..7

Fig. 3: Description of the different programmable nucleases and three major repair pathways……… 9

Fig. 4: Schematic representation of DNA binding by zinc finger protein………...10

Fig. 5: Schematic representation of DNA binding by TAL effector………...12

Fig. 6: Three stages of CRISPR immunity: adaption, crRNA biogenesis, and interference...14

Fig. 7: CRISPR-Cas9 mediated DNA double strand break………...15

Fig. 8: Homology-directed DNA repair pathway………....20

Fig. 9: Non homologous end joining DNA repair pathway………....21

Fig. 10: Microhomology-mediated end joining DNA repair pathway………..23

Fig. 11: Design of currently available templates………...26

Fig. 12: The traffic light reporter system………...28

Fig. 13: Optimizing DNA double strand break repair in modified TLR3 system………….…47

Fig. 14: TLR1, TLR2, and TLR3 system………..48

Fig. 15: Cloning strategy of TLR3………49

Fig. 16: NHEJ and HDR controls of TLR1, TLR2, and TLR3 system……….50

Fig. 17: Comparison of TLR and TLR-delTA sequence………...51

Fig. 18: Microscopy and FACS analysis of TLR control plasmids…...………...52

Fig. 19: CRISPR-Cas9 target sites………54

Fig. 20: Two CRISPR-Cas9 expression systems………..55

Fig. 21: Activity test of both expression systems……….56

Fig. 22: Comparison of two CRISPR-Cas9 systems………57

Fig. 23: Generation of different donor templates……….58

Fig. 24: Gel analysis of donor templates………..59

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Fig. 26: Schematic representation of the generation of HEK 293-TLR3 stable cell line……..61

Fig. 27: Gel analysis of TLR expression cassette with CMV promoter and BGH polyA signal……….…62

Fig. 28: Activity test of px459 CRISPR-Cas targets in HEK-TLR3 stable cell line……….…63

Fig. 29: Activity test of gRNA cloning vector and hCas9 expression vector in HEK-TLR3 stable cell line………...64

Fig. 30: px459-mRFP variant of CRISPR-Cas9 system………....…65

Fig. 31: Optimizing HDR events using different donor templates………66

Fig. 32: Optimizing the HDR events using NHEJ key protein inhibitor………...67

Fig. 33: Interaction of Cas9 with target DNA………73

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List of tables

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List of tables

Table 1: Chemicals and reagents……….……….30

Table 2: Buffers……….………...31

Table 3: Media……….……….32

Table 4: Plasmids and expression vectors……….…………...32

Table 5: Oligonucleotides……….………33

Table 6: Restriction enzymes………...36

Table 7: Polymerases………37

Table 8: Markers……….………..37

Table 9: DNA purification kits……….………37

Table 10: DNA cloning kits………...38

Table 11: Bacterial strains………...…………..38

Table 12: Devices………...38

Table 13: PCR approach……….40

Table 14: PCR program………..41

Table 15: gRNA sequences targeted TLR3 system………53

Table 16: Comparison of assays for quantifying genome editing outcomes….……….69

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1. Introduction

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1. Introduction

1.1 Gene Therapy

Gene therapy is defined as the transfer of nucleic acids (DNA or RNA) to patients’ cells for the treatment of human diseases. It can be divided into two categories: “in vivo” and “ex vivo” gene therapy. In the first approach, the therapeutic gene is directly introduced into the patients’ body (e.g. muscle, liver), while in the second approach, the patients’ cells are isolated from the body, genetically modified in the laboratory and reintroduced into the patients’ body.

Depending on the type of disease, gene therapy can be achieved either by addition of a functional cDNA copy as a substitute for the mutated gene to restore the normal genetic function (gene addition) or by using RNA interference to knock down the dominant negative and toxic gain-of-function gene products (gene silencing). Recently, gene targeting therapy has been brought to the forefront as a potential therapy approach for many monogenetic diseases, which aims at correcting the defective endogenous counterpart through homologous recombination based DNA double strand break repair (DSBR). Besides the correction of mutations that cause diseases, this genome-editing technology also enables the scientist to add therapeutic genes to the specific site in the genome and to precisely remove the mutated genome sequence. These innovative approaches have generated great enthusiasm and successfully moved from bench to bedside (Kaufmann, Buning et al. 2013).

1.1.1 Gene therapy in the past

In 1990, the first “ex vivo” gene therapy was performed by Dr. William French Anderson on two children with adenosine deaminase deficiency (ADA-SCID), a monogenetic disease leading to severe combined immunodeficiency. The T cells were isolated from these two patients and

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transduced with an ADA-containing retroviral vector to express normal ADA gene. Both patients have shown increased T lymphocyte counts in the blood as well as ADA enzyme activity in one patient after the treatment (Blaese, Culver et al. 1995). Although the effects were temporary and many components remain to be perfected, it sets a new milestone in the development of gene therapy.

With this notable success, more promising results have been obtained from the clinical trials suffering from Leber’s congenital amaurosis (Bainbridge, Smith et al. 2008, Maguire, Simonelli et al. 2008), haemophilia (Lheriteau, Davidoff et al. 2015), β-thalassemia (Cavazzana-Calvo, Payen et al. 2010), diabetes (Elsner, Terbish et al. 2012) and Parkinson’s disease (Stoessl 2014). Between 1990 and 2013 more than 1900 clinical trials have been conducted with a varying degree of success and no major side effects reported (Kaufmann, Buning et al. 2013).

In 2004, China introduced the world’s first commercial gene therapy drug (Gendicine®

) into the market for the treatment of patient with head and neck squamous cell carcinoma (HNSCC) (Pearson, Jia et al. 2004, Peng 2005,Wilson 2005). Gendicine® is a recombinant human serotype 5 adenovirus genetically engineered to express the human p53 gene. The p53 gene is one of the most widely studied tumor suppressor genes, which plays a crucial role in preventing cancer formation. Because of its important role in preventing genome mutation and maintaining genome stability, the p53 gene has been described as "the guardian of the genome". After intratumoral injection of Gendicine®, the adenoviral particle delivers the therapeutic p53 gene to the cytoplasm and nucleus for the expression without integrating it in the host cells’ chromosomes. In clinical trials and application, infiltration of many lymphocytes and inhibition of VEGF activity in biopsies of tumor lesions were observed. The combination of Gendicine® with radiotherapy has shown significant synergistic effects in Phase II/III clinical trials, with 64% complete regression and 29% partial regression (Peng 2005). The only side effect of Gendicine® is self-limited fever after more than five years clinical observation (Pearson, Jia et al. 2004). Gendicine® represents a remarkable medical achievement, and it opened the door for a gene therapy market in the near future.

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In Europe, the first commercial gene therapeutic product Glybera® was approved by the European Commission at the end of 2012 (Miller 2012). Glybera® is an adeno-associated viral vector engineered to express lipoprotein lipase (LPL) for the treatment of familial lipoprotein lipase deficiency (LPLD). Familial LPLD represents a rare autosomal recessive disorder usually present in childhood characterized by severe hypertriglyceridemia with abdominal pain, recurrent acute pancreatitis, eruptive cutaneous xanthomata, hepatosplenomegaly, and other complications (Brunzell 1993). Familial LPLD is caused by extremely low or absent activity of LPL, the key enzyme responsible for hydrolysis of triglyceride-rich lipoproteins (Goldberg 1996). LPL interacts with circulating chylomicrons in the vascular lumen and converts triglyceride into free fatty acids (Bryant, Christopher et al. 2013). Without LPL, triglyceride cannot be depleted, leading to the accumulation of triglyceride-rich lipoproteins in the plasma (Goldberg 1996). After injection of Glybera® into the leg muscle, patients showed a long-term reduction of triglyceride levels (Gaudet, Methot et al. 2012). Although the clinical results were based on a small number of patients, the marketing authorization for Glybera® represents a landmark in the gene and cell therapy field.

Since the successes in the gene therapy area are based on the addition of a therapeutic gene for the treatment of loss of function genetic diseases, the application is very limited by other types of diseases like autosomal recessive and gain-of-function diseases. Therefore, great efforts are made in the genome editing field to cut and repair the endogens precisely though homologous recombination, which can be applied to any kind of monogenic diseases.

1.1.2 Gene transfer

Despite clinical success, the understanding of various gene transfer tools and molecular mechanisms has resulted in the development of gene therapy approaches with improved safety and therapeutic efficacy. Over the past years, genetically engineered viral and non-viral vectors are widely used for the delivery of therapeutic genes to the specific human cells.

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Viral vectors, such as adenoviral vectors, adeno-associated viral (AAV) vectors, lentiviral vectors, and retroviral vectors, are the most common viral delivery vectors, which provide efficient gene transduction and effective gene expression in over 68% of gene therapy clinical trials (Santiago-Ortiz and Schaffer 2016).

AAV vectors in particular are widely used for many in vivo gene transfer applications to both dividing and non-dividing cell populations with low host immune response, high transferring ability, long-term gene expression, and low toxicity (Kotterman and Schaffer 2014). As a result of these properties, AAV is becoming a promising approach to treat a variety of diseases and cancers, including hemophilia B, LPLD, Alzheimer’s disease, Parkinson’s disease, inherited retinal dystrophies (IRD), and liver cancer (Luo, Luo et al. 2015).

AAV is a linear single-stranded DNA (ssDNA) parvovirus. The wild-type AAV genome, which is about 4.7 kb long, comprises two open reading frames encoding Rep and Cap protein flanked by two hairpin palindromic repeat sequences termed inverted terminal repeats (ITRs) (Santiago-Ortiz and Schaffer 2016). The Rep gene codes for non-structural proteins expressed via alternative promoters, which are involved in viral replication, transcription, packaging, and genomic integration. The Cap gene encodes the structural proteins VP1, VP2, VP3 that assemble to form the viral capsid (Deyle and Russell 2009). In the absence of a helper virus, the AAV genomes can establish latency and persist in the host as episomes or in some cases integrate into the host genome on the long arm of chromosome 19 in a site-specific manner. In the presence of a helper virus such as adenovirus or herpesvirus, the AAV can enter its replication cycle and undergoes productive infection (Deyle and Russell 2009). Infection is initiated by low-affinity binding to glycans followed by cell surface receptor-mediated endocytosis. Endosomal escape, endosomal trafficking, and viral capsid uncoating occurs and the single stranded DNA undergoes double strand synthesis, which is capable of transcription and gene expression(Lisowski, Tay et al. 2015).

Recombinant adeno-associated viruses (rAAVs) are generated by replacing the viral ORFs with transgene expression cassettes containing the gene of interest. Rep and Cap genes, as well as

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helper genes required for AAV replication are provided in trans (Fig. 1). Either single-stranded DNA genomes of 4.7 kb or self-complementary DNA genomes of approximately 2.2 kb can be packed into AAV vectors (Gray and Zolotukhin 2011). For large therapeutic genes (> 4.7 kb), different strategies have been developed to expand the packaging capacity imposed by the viral genome. These include minimizing the expression elements in the expression cassette, truncating the gene itself, or using trans-splicing and overlapping vectors based on post-transduction concatemerization of the transgene (Chira, Jackson et al. 2015).

Fig. 1: Generation of recombinant AAV vectors. Recombinant AAV vectors are generated by replacing the

viral ORFs with transgene expression cassettes containing the promoter, therapeutic gene and pA sequence. At the left side of the picture is the hexagonal three-dimensional structure of the viral capsid and electron microscopy image of the individual 21nm large virus (Stieger and Lorenz 2008).

However, their applications are limited to the inherent proprieties of the target gene. A hybrid dual vector system has been developed as a potential solution to this problem by inserting a

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highly recombinogenic alkaline phosphatase (AP) sequence in the trans-splicing vector, which allows the AP sequence-mediated transgene reconstitution through homologous recombination (Ghosh, Yue et al. 2008). These recent advances may offer a strong potential for large gene reconstitution, however, further understanding of vector-host-interaction and endogenous mechanisms are required for achieving successful therapeutic application.

Another important type of vector for gene therapy is derived from lentiviruses (LV), which have been used particularly for the treatment of central nervous system disorders. Lentiviruses are able to transduce both dividing and non-dividing cells with high transduction efficacy, long-term stable expression of the transgene, and low immunogenicity (Chira, Jackson et al. 2015). Typically, vector particles are produced by cotransfection of the lentiviral vector plasmid and three helper constructs (pMDL, pRev and pVSVG) in the packaging cell. The vector plasmid contains the vector genome and the transgene, while the helper constructs encode the proteins that are essential for the viral life cycle (Tiscornia, Singer et al. 2006). Compared to AAVs, LV can accommodate larger transgenes up to 10 kb, which broadens the applicability of LVs for gene therapy application (Matrai, Chuah et al. 2010) (Fig. 2).

Upon transduction of the target cells, the virion enters the host cells by binding to the cell surface receptors and co-receptors, leading to endocytosis or direct fusion with the cell membrane. After entry, the internal core is released and the RNA transgene is reverse transcribed to the cDNA. The viral core containing the cDNA is then transported to the nucleus and the cDNA is integrated into the host cell chromosome, resulting in persistent expression of the transgene (Tang, Kuhen et al. 1999). However, activation of proto-oncogenes at the site of genomic integration suggests the carcinogenic effect of LVs in the host genome. To overcome this limitation, self-inactivating (SIN) vectors and integration-deficient lentiviral vectors (IDLV) have been developed by deleting the U3 region of the 3’LTR and mutational inactivation of the integrase protein, respectively (Matrai, Chuah et al. 2010). These recent achievements highlight the improved safety of LVs with important implication for further clinical purposes.

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Fig. 2: Recombinant lentivirus vectors. The viral ORFs of the wild type Lentiviruses are replaced by the

expression cassette of the therapeutic gene, which is maximal 7 kb long (Stieger and Lorenz 2014). LTR: Long terminal repeat; SIN: Self inactivating LTR; RRE: Rev Response Element; Ψ: Pack sequence psi.

Despite accumulating data on improved viral vectors, non-viral vectors, such as naked plasmids, nanoparticles, cationic liposomes and cationic polymers have also shown their advantages in the gene therapy field. In contrast to viral-vectors, non-viral vectors are typically easy to synthesize, and the immunogenicity is lower. Non-viral vectors are also capable of delivering large therapeutic genes and synthetic expression cassettes like siRNA. The limitations of non-viral vectors are related to extracellular stability, internalization, intracellular trafficking, nuclear entry, and the sustainability of expression of the transgene. Several strategies have been developed to overcome these limitations by using carrier molecules, targeting ligands, endosomal disruptive agents, and nuclear localization signal (Ramamoorth and Narvekar 2015).

Besides these recent improvements in gene delivery vectors and a large amount of clinical successful events, the development of programmable nucleases and the understanding of nuclease functions opened the new era of genome editing field.

1.2 Highly specific nucleases

In recent years, several programmable nucleases like meganucleases, zinc-finger nucleases (ZNFs), transcription activator-like effector nucleases (TALENs) and RNA-guided

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endonucleases (RGNs) have become powerful tools for precise and efficient genome engineering. These engineered nucleases can create site-specific DNA double-strand breaks, and the induced double-strand breaks can be repaired through different DNA repair pathways including homologous recombination, non-homologous end joining, and microhomology-mediated end joining (Fig. 3). To induce site-specific DNA DSB in the human genome of 3 x 109bp, the recognition site of the nucleases should be 16-18 bp in length (Chandrasegaran and Carroll 2016).

To date, ZNFs and TALENs have been used in more than 40 different organisms and cell types, which have shown their successes in genome editing area (Chandrasegaran and Carroll 2016). A more recent genome editing tool is the CRISPR-Cas9 system, which has been described as an adaptive defense mechanism in bacteria and archaea. In the past three years, CRISPR-Cas systems have been used in a wide range of organisms, including Drosophila melanogaster (Gratz, Cummings et al. 2013), Caenorhabditis elegans (Friedland, Tzur et al. 2013), Saccharomyces

cerevisiae (DiCarlo, Norville et al. 2013), zebrafish (Chang, Sun et al. 2013), mice (Wang, Yang

et al. 2013), rat (Li, Qiu et al. 2013, Li, Teng et al. 2013), plants (Jiang, Zhou et al. 2013), monkeys (Niu, Shen et al. 2014) and human embryos (Kang, He et al. 2016).

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Fig. 3: Description of the different programmable nucleases and three major repair pathways. (A)

Schematic drawing of ZNF, TALEN, and CRISPR-Cas. The DNA binding domains are represented in turquoise, the cleavage domains are in yellow. (B) Three major repair pathways occur after the introduction of the DNA double strand breaks, including NHEJ, HDR, and MMEJ (Yanik, Muller et al. 2016).

1.2.1 Zinc finger nucleases

ZNFs were the first artificial targetable nucleases, which can be customized to cleave any given sequence in the genome (Kim and Berg 1996). ZFNs consist of a programmable DNA-binding domain of zinc finger proteins (ZFPs) and a non-specific DNA cleavage domain of type II restriction endonuclease FokI (Kim, Cha et al. 1996). Because the binding domain of the ZFPs can be easily manipulated, they have become powerful tools for genome engineering and have

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been used to site-specifically modify the genomes in various organisms, including frogs, insects, fish, plants, mice, rats and cultured human cells (Wu, Kandavelou et al. 2007). Furthermore, ZFNs are the only genome editing tools that have been tested in clinical trials for the treatment of Hemophilia B, Mucopolysaccharidosis I and HIV infections (Yanik, Muller et al. 2016).

Fig. 4: Schematic representation of DNA binding by zinc finger protein. Each zinc finger usually

recognizes 3 base pairs DNA through four contact amino acids at positions 1, 2, 3, 6 of each helix (shown in purple) (Pingoud, Wilson et al. 2014).

The DNA-binding domain of ZFNs contains 3 to 6 Cys2-His2 zinc fingers. Each zinc finger (ZF)

is composed of approximately 30 amino acid residues in a unique ββα configuration, which is stabilized by a zinc atom (Pavletich and Pabo 1991). An individual zinc finger usually recognizes 3 base pair DNA sequences by inserting an α-helix into the major groove of the DNA double helix (Pavletich and Pabo 1991) (Fig. 4). The binding specificity of the ZFP has a direct influence on the cleavage specificity, which can be achieved by adding more ZF motifs to the ZFPs, but the recognition of DNA by individual ZFs appears to be dependent of the neighboring modules (Urnov, Miller et al. 2005, Urnov, Rebar et al. 2010). To construct multi-finger arrays and to improve the binding specificity, different strategies have been developed, including OPEN (Oligomerized Pool ENgineering) and CoDA (Context-Dependent Assembly) (Maeder, Thibodeau-Beganny et al. 2008, Sander, Dahlborg et al. 2011).

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To produce a DSB, the nuclease domain of the FokI endonuclease must be dimerized for the formation of the active center (Bitinaite, Wah et al. 1998, Wah, Bitinaite et al. 1998). Therefore, a pair of ZFNs binding to adjacent, oppositely oriented sites on the DNA is required for the induction of the DSB (Smith, Bibikova et al. 2000). Paired binding sites of ZFNs doubled the size of the target sequence recognition and increased the specificity of ZFNs. Normally, a pair of 3- or 4- finger ZFN monomers has an 18- or 24- bp recognition site in a tail-to tail orientation, which is long enough to specify a unique genomic sequence in mammals and plants (Wu, Kandavelou et al. 2007). However, due to the unspecific DNA cleavage domain and unexpected binding to nonspecific DNA sequences, significant off-target cleavage effects have been observed besides the on-target cleavage, indicating that still great efforts should be made to improve the specificity of the ZFNs (Gabriel, Lombardo et al. 2011, Pattanayak, Ramirez et al. 2011).

1.2.2 TALENs (transcription activator-like effector nucleases)

TALENs (transcription activator-like effector nucleases) are another type of highly specific nucleases, which can be designed to cleave any given DNA sequences for targeted modification of endogenous genes. The general modular structure of TALENs is similar to that of ZFNs, which is based on a specific DNA binding domain of bacterial TALEs fused to the unspecific DNA cleavage domain.

The DNA binding domains TALEs (transcription activator-like effectors) are transcriptional activators that are derived from the bacterial plant pathogen Xanthomonas (Bonas, Stall et al. 1989). TALEs contain N- and C- termini for localization and a central repeat domain for specific DNA binding (Boch and Bonas 2010). The central repeat domain of TALE proteins is composed of 5 to over 30 tandem repeats with an average of 17.5 (Wei, Liu et al. 2013). Each repeat contains 33-35 amino acid residues and recognizes one base pair in the target DNA via unique repeat variable di-residues (RVD) at position 12 and 13 (amino acid residue NI recognizes A, HD recognizes C, NG or HG recognizes T, and NN recognizes G or A), which determines the nucleotide binding specificity of each repeat (Deng, Yan et al. 2012, Mak, Bradley et al. 2012)

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(Fig. 5). Since there is no considerable context-dependent interaction between the neighboring

repeats, TALENs have much more advantages compared to ZFNs (Yanik, Muller et al. 2016).

Fig. 5: Schematic representation of DNA binding by TAL effector.The DNA binding domain of

TALEs consists of a series of repeats and each repeat recognizes one base pair in the target DNA via repeat variable di-residues (RVD) at position 12 and 13. The RVD (HD) is shown in red (Pingoud, Wilson et al. 2014).

By fusing the nuclease cleavage domain such as FokI and PvuII with an artificial TALE binding domain, TALENs have been used in a wide range of model organisms and cell types, including flies, frogs, fish, rats, mice, human somatic cells, and human cells (Miller, Tan et al. 2011, Wei, Liu et al. 2013, Yanik, Muller et al. 2016). In 2015, the first-in-man application of TALEN engineered universal CAR19 T cells took place and will now be tested in clinical trials (Yanik, Muller et al. 2016).

1.2.3 CRISPR-Cas system

Recently, the development of CRISPR-Cas (clustered regularly interspaced short palindromic repeats-CRISPR-associated proteins) systems is revolutionizing the field of genome editing, enabling the scientists to manipulate the genomes for therapeutic application with relative ease.

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The CRISPR-Cas systems are prokaryotic immune systems, which provide adaptive immunity against invading phages and foreign nucleic acids. CRISPR-Cas systems are present in roughly half of all sequenced bacterial genomes and almost all of sequenced archaeal genomes (Barrangou, Fremaux et al. 2007, Wright, Nunez et al. 2016).

In these species, CRISPR arrays contain a series of short, palindromic DNA repeats ranging from 21 to 48 bp, interspaced by 26 to 72 bp variable spacer sequences derived from invading pathogens (Bondy-Denomy and Davidson 2014). The CRISPR array is usually located adjacent to a cluster of CRISPR-associated (cas) genes and preceded by an AT-rich leader sequence (Amitai and Sorek 2016). Depending on the Cas genes and the proteins they encode, three major types of CRISPR-Cas systems have been identified, namely Type I, Type II and Type III, and these can be further divided into several subtypes, given their structural and functional diversity (Amitai and Sorek 2016). The key protein of Type I systems is Cas3, while the signature protein of Type III systems is Cas10. The most widely used CRISPR systems are the Type II CRISPR-Cas9 systems from Streptococcus pyogenes, which are signified by CRISPR-Cas9 protein (Wright, Nunez et al. 2016).

Overall, CRISPR immunity can be divided into three stages: adaption, CRISPR RNA (crRNA) biogenesis, and interference (van der Oost, Westra et al. 2014, Makarova, Wolf et al. 2015). In the adaption stage, foreign DNA is identified and integrated into the CRISPR array as a new spacer. During the crRNA biogenesis stage, the CRISPR array is transcribed and processed into small crRNAs that each contains a single spacer flanked by CRISPR repeat sequences. These mature crRNAs are subsequently combined with Cas proteins for the formation of the active Cas-crRNA complex. In the interference stage, the Cas-Cas-crRNA complex recognizes foreign nucleic acid sequence complementarity to the crRNA sequence, which leads to the successful cleavage and degradation of the DNA and RNA molecules (Fig. 6).

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Fig. 6: Three stages of CRISPR immunity: adaption, crRNA biogenesis, and interference. During the

adaption stage, foreign DNA is identified and integrated into the CRISPR array as a new spacer. In the crRNA biogenesis stage, the CRISPR array is transcribed and processed into small crRNA, which combines with Cas protein and forms the active Cas-crRNA complex. The Cas-crRNA complex recognizes and cleaves the foreign DNA 3 bp upstream of the PAM sequence.

The engineered Type II CRISPR-Cas9 system consists of the Cas9 DNA endonuclease and a chimeric single guide RNA (gRNA). The gRNA contains a 20 nt programmable CRISPR RNA (crRNA) and a trans-activating crRNA (tracrRNA), which forms a double-stranded RNA structure and binds to Cas9that form a complex for cleavage (Jinek, Chylinski et al. 2012, Hsu, Lander et al. 2014). Next to the target sequence, a 2-5 nucleotide motif is required for the target recognition; named protospacer adjacent motif (PAM) that usually consists of a 5’-NGG-3’ trinucleotide, in which N can be any nucleotide (Horvath and Barrangou 2010). Upon recognition

Adaption

crRNA biogenesis

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of gRNA and target DNA sequence, the Cas9-gRNA-target DNA ternary complex initiates the subsequent cleavage of the target strand by the HNH nuclease domain, and of the non-target strand by the RuvC domain (Fig. 7) (Amitai and Sorek 2016).

Fig. 7: CRISPR-Cas9 mediated DNA double strand break. (A and B) Crystal structure of

Cas9-sgRNA-dsDNA ternary complex. The target DNA strand and nontarget strand are colored dark blue and purple, respectively. sgRNA is shown in orange (Amitai and Sorek 2016) (C) Upon the recognition of 20 bp target DNA next to the PAM sequence, DNA double strand break is mediated by the two activate centers of the Cas9 protein, RuvC and HNH. TS, target strand; NTS, nontarget strand.

A

B

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Beyond the introduction of the DNA breaks, CRISPR technologies have also been used in different ways, such as regulating gene expression, modifying epigenomes, and dynamic imaging of chromatin. Specifically, the CRISPR-associated catalytically inactive dCas9 protein has been fused to transcription repressor or activator domains for the regulation of gene expression in human and yeast cells (Gilbert, Larson et al. 2013, Larson, Gilbert et al. 2013,Gilbert, Horlbeck et al. 2014). By using eGFP-tagged dCas9 protein and sequence-specific gRNA, dCas9 chimeras enable the imaging of DNA and visualization of chromatin organization and dynamics in living human cells (Chen, Gilbert et al. 2013). Likewise, the CRISPRainbow technique based on dCas9 combined with fluorescence-labelled gRNA has demonstrated simultaneous imaging of up to six distinct chromosomal loci in living cells (Ma, Tu et al. 2016). Recently, CRISPR-Cas9-based acetyltransferases and demethylases enable the epigenetic regulation and provide a new tool for manipulating gene expression (Hilton, D'Ippolito et al. 2015, Pham, Kearns et al. 2016).

1.3 DNA double strand break repair

1.3.1 DNA damage responses (DDR)

In human cells, DNA damage takes place at a rate of 10,000 to 1,000,000 molecular lesions per cell per day. It can be caused by exogenous agents and endogenous cellular processes, such as ionizing radiation (IR), ultraviolet light (UV), chemical agents, and replication errors (Hoeijmakers 2001, Hoeijmakers 2001). One of the most dangerous types of DNA damage is a double-strand break (DSB) that can result in the introduction of gene mutations, chromosome rearrangement, and cell death (Khanna, Lavin et al. 2001). DNA double strand breaks can also be induced artificially by using highly specific nucleases and through the addition of template DNA to trigger the desired repair outcomes. Efficient and accurate DNA repair is crucial for the maintenance of genomic stability and prevention of tumor formation. Designed DNA cut and repair can be used as a potential therapeutic approach to repair the disease causing mutations. In mammalian cells, DNA double-strand breaks (DSBs) can be repaired by different pathways,

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homologous recombination (HR), classical nonhomologous end joining (c-NHEJ), microhomology-mediated end joining (MMEJ), and single-strand annealing (SSA) (Helleday, Petermann et al. 2008). In the cell division cycle, multiple cyclin-dependent kinases (CDKs) are periodically activated and play a central role in DNA repair pathway choices. C-NHEJ can occur in any phase of the cell cycle but is dominant in G0/G1 and G2, whereas HR usually takes place in S and G2 cell phases because it uses sister-chromatid sequences as the template for repair (Sancar, Lindsey-Boltz et al. 2004).

In addition to these repair pathways, DNA damage response (DDR) also plays a key role in combating threats posed by DNA damage. It is a signal transduction pathway that enables the cell to detect DNA lesions, propagate DNA damage signals, and promote their repair. DDR pathway is mediated by the activation of ataxia-telangiectasia mutated (ATM), ataxia-telangiectasia RAD3-related kinase (ATR), and DNA-dependent protein kinase (DNA-PK). ATM is primarily activated by double-stranded DNA breaks (DSBs), whereas ATR responds to RPA-coated single-stranded DNA (ssDNA) region.

The key regulator of ATM activation is the Mre11-Rad50-Nbs1 (MRN) complex, which functions as a DSB sensor and is required to recruit DDR downstream proteins. DNA-PK promotes DSB religation and is involved in the non-homologous end joining pathway of DNA repair (Goldstein and Kastan 2015). Once the activated ATM and ATR kinases are at the DSB, they phosphorylate a number of substrates such as H2AX, NBS1, CHK1, BRCA1, p53, and CHK2. The phosphorylated form of H2AX (known as γH2AX), is recognized by the mediator of DNA damage checkpoint 1 (MDC1), which then recruits ring finger protein 8 (RNF8), an E3 ubiquitin ligase. RNF8 promotes another E3 ligase ring finger protein 168 (RNF168) to ubiquitinate H2A-type histones, leading to the recruitment of the p53 binding protein 1 (53BP1) and receptor-associated protein 80 (RAP80) to DSB sites (Bohgaki, Bohgaki et al. 2013).

After the DNA damage responses, different repair key proteins affect the decision of the repair pathway choices with collaboration and competition (Kass and Jasin 2010). Understanding of the

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repair mechanisms helps us to inhibit the error-prone repair pathway and bias the repair outcomes toward HDR.

1.3.2 Homologous recombination (HR)

HR is a central pathway for accurate DNA double strand break repair (DSBR), which is described as an error-free repair mechanism. HR uses an undamaged homologous sequence as a donor template for repair and requires RAD51-mediated strand invasion. HR is initiated by resection of DNA ends at the DSB site. In most cases, DNA end resection in eukaryotes is a two-step process (Mimitou and Symington 2008, Zhu, Chung et al. 2008,Ceccaldi, Rondinelli et al. 2016). In the initial phase of the end resection, a small number of base pairs (fewer than 20 bp in mammalian cells, 100-200 bp in yeast) are processed by MRE11-RAD50-NBS1 (MRN) complex and CtBP-interacting protein (CtIP) (Zhu, Chung et al. 2008, Truong, Li et al. 2014).

Following the initial DNA processing, the extension resection (which is known as 5’-3’ resection) is mediated by helicases and nucleases (i.e., CtIP, EXO1, DNA2, BLM, WRN) to generate a long stretch of 3’ single-strand DNA (ssDNA) for strand invasion (Sturzenegger, Burdova et al. 2014, Ceccaldi, Rondinelli et al. 2016). During this process, end resection is promoted by cyclin-dependent kinase (CDK)-cyclin-dependent phosphorylation of multiple substrates, such as DNA2, CtIP, and EXO1 (Yun and Hiom 2009, Chen, Niu et al. 2011,Tomimatsu, Mukherjee et al. 2014). Next, the resected DNA is coated by ssDNA-binding protein replication protein A (RPA) to minimize the formation of secondary structures. RPA is then displaced and the Rad51 is loaded onto the ssDNA to form a nucleoprotein filament, a step that is mediated by BRCA2 (breast cancer type 2) and RAD52 (Renkawitz, Lademann et al. 2014). After RAD51 formation, homology search and DNA strand invasion takes place leading to D loop formation between the broken DNA and the intact homologous donor sequence (Sancar, Lindsey-Boltz et al. 2004).

More than three different pathways are proposed after the D-loop intermediate. In the classic double-strand break repair (DSBR) model, the 3’ end in the D loop is extended by repair

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synthesis and the second DSB end aligns with the extended D‑loop to form a double holliday junction (DHJ). Crossover and non-crossover overcomes are produced by resolvases or combined helicase/topoisomerase, such as GEN1, MUS81-EME1 complex, and BLM-TOPOIIIα-RMI1-RMI2 (BTR) complex (Matos and West 2014). According to the synthesis-dependent strand annealing (SDSA) model, the nascent strand from the D-loop anneals to the 3’ end of the broken chromosome and the single-stranded gaps are filled in by DNA synthesis and ligation. This type of repair results in non-crossover products (Rodgers and McVey 2016). In break-induced replication (BIR), only one end of the DSB aligns homology with the template and the D-loop is assembled into the replication fork (Fig. 8). Replication of the entire homologous template arm results in a large-scale loss-of heterozygosity (LOH) (Pardo, Gomez-Gonzalez et al. 2009).

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Fig. 8: Homology-directed DNA repair pathway. HDR is initiated by DNA end resection at the DSB

site, followed by the binding of RPA and Rad51. In the presence of DNA donor template, precise DNA repair takes place resulted in crossover or non crossover products.

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1.3.3 Non homologous end joining (NHEJ)

NHEJ has been considered as error-prone repair of DSBs, which is characterized by re-ligating of two broken ends independently of sequence homology. NHEJ is a predominant DNA double strand repair pathway, which is always associated with the introduction of small insertions and deletions (indels) at the break site. Non homologous end joining is initiated by the binding of the KU70/80 heterodimer, followed by recruitment and activation of the DNA protein kinase catalytic subunit (DNA-PKcs). Binding of the Ku heterodimer protects the broken ends from extensive resection and inhibits their degradation. DNA-PKcs undergoes autophosphorylation and phosphorylates other NHEJ downstream proteins, such as X-ray repair cross-complementing protein 4 (XRCC4) and Artemis. DNA ends are subsequently re-ligated by the XRCC4-ligase IV-XLF complex (Hoeijmakers 2001, Helleday, Petermann et al. 2008) (Fig. 9).

Fig. 9: Non homologous end joining DNA repair pathway. After the DNA DSBs induced

by specific nucleases, KU70/80, DNA-PKcs, XRCC4, and Artemis bind at the break site. The ends are finally re-ligated by XRCC4-ligase IV-XLF complex, which always results indels.

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1.3.4 Microhomology-mediated end joining (MMEJ)

In the absence of c-NHEJ factors such as Ku70, Ku80, or DNA ligase IV, yeast and mammalian cells are still able to repair DSBs via an alternative form of DSBR, termed microhomology-mediated end joining (MMEJ) (Deriano and Roth 2013). MMEJ is an error-prone repair mechanism that always generates small deletions flanking the DSBs. MMEJ is also associated with chromosomal translocations and telomere fusions, thereby resulting in harmful consequences on genomic stability (McVey and Lee 2008).

In mammalian cells, both MMEJ and HR share the common initial end resection step mediated by MRN complex and CtIP in a CDK-dependent manner, which reveals 5-25 base pair (bp), single-strand microhomologous sequences for further sequence alignment (McVey and Lee 2008). Repair is completed by annealing of microhomologies, cleavage of 3’ flaps, fill-in DNA synthesis, and ligation. Although the MMEJ repair mechanism is still less characterized, numerous studies highlight critical roles for XRCC1/DNA ligase III complex, PARP1 and translesion synthesis (TLS) DNA polymerase theta (polθ) in regulating MMEJ in higher organisms (Sfeir and Symington 2015) (Fig. 10).

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Fig. 10: Microhomology-mediated end joining DNA repair pathway. In

the presence of microhomology region in the DNA, the DSBs can also be repaired via MMEJ. PARP1, MRE 11 complex, and CtIP plays an important role in MMEJ DNA repair.

1.3.5 DNA donor templates

The efficiency of HDR can be influenced by a lot of endogenous and exogenous factors, including the cell cycle, the cell type, chromosomal region, the activity of the repair system, and the DNA donor template (Yanik, Muller et al. 2016). To enhance HDR, different approaches have been developed, including the manipulation of the cell cycle and the regulation of expression of key repair pathway proteins (Chu, Weber et al. 2015, Maruyama, Dougan et al. 2015,Srivastava and Raghavan 2015). However, these invasive manipulations may be undesirable for therapeutic applications because they can alter the cellular response to DNA damage at other non-target sites in the genome and lead to tumor formation.

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In contrast, designing optimal DNA donor templates can increase HDR frequencies and at the same time leave cell cycle regulation untouched. Linearized or double-stranded DNA plasmid sequences, as well as ssDNA oligonucleotides, are used as template for homologous recombination at the target site (Fig. 11). Viral vectors such as AAV or IDLV can also be used as a source of donor DNA, which provide single-stranded DNA as template for HDR (Hirsch, Green et al. 2010, Handel, Gellhaus et al. 2012, Coluccio, Miselli et al. 2013, Genovese, Schiroli et al. 2014).The size of the intended sequence changes, the length of the homology arms, and the insertion site of the mutation are important factors to be considered. Although the exact mechanism by which donor design increases HDR frequencies is still under investigation, several evidences have shown its influence on gene targeting outcomes.

In mammalian cells, a plasmid donor with at least 1-2 kb of total homology is usually used for creating large sequence changes in the presence of target cleavage, including insertion of reporter genes such as fluorescent protein or antibiotic resistance markers (Dickinson, Ward et al. 2013, Yang, Wang et al. 2013). Without target cleavage, a total of 8-15 kb homology is normally used (Wu, Ying et al. 2008). Generally, the efficiency of recombination increases as the length of homology arms increases, while the efficiency decreases as the size of the DNA insert increases (Li, Wang et al. 2014), but if the homology arms contain repetitive DNA sequences, the targeting efficiency will be low (Wu, Ying et al. 2008).

In many proof-of-concept studies, it has been demonstrated that disease-causing mutation can be corrected by using DNA donor templates along with targeted nucleases. For the correction of the

IL2Rγ gene mutation, ca. 1,543 bp centered plasmid donor has been used together with ZFN,

which has shown about 7% of cells with desired genetic modification (Urnov, Miller et al. 2005).

For small sequence changes, ssDNA sequences are usually more efficient than plasmid donors. To correct the duchenne muscular dystrophy gene (Dmd) mutation in the germ line of mdx mice, a single-stranded oligodeoxynucleo-tide (ssODN) has been used as a template for HDR-mediated gene repair, which contains 90 base pairs (bp) of homology sequence flanking each side of the target site (Long, McAnally et al. 2014). Four single stranded Crb1rd8 correction ssODNs

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mer and 52-mer, in sense and antisense directions) with homology centered to the targeted region has been compared to stimulate HDR events for correction of the Crb1rd8 allele in C57BL/6N mice, in which 200-mer sense ssODN has shown the best result (Low, Krebs et al. 2014).

Recently, enhanced HDR rates have been reported by using optimized asymmetric ssDNA donor templates for conversion of a BFP reporter gene into a GFP reporter gene via mutation of three nucleotides within the BFP reading frame (Richardson, Ray et al. 2016). In this study, it has been observed that donor DNA complementary to the nontarget strand is more effective than donor complementary to the target strand. The optimized donor DNA is complementary to the nontarget strand by overlapping the Cas9 cut site with 36 bp in the PAM-distal side, and with 91 bp on the PAM-proximal side (Richardson, Ray et al. 2016). It was shown that optimizing a donor template at the 5’ and 3’ homology regions flanking the DSB site could boost the frequency of HDR in the absence of chemical and genetic intervention.

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Fig. 11: Design of currently available templates. Templates can be generated as double stranded plasmid,

linearized plasmid, PCR product, single stranded (ss)DNA or viral vector DNA (AAV or IDLV) (Yanik, Muller et al. 2016).

1.3.6 Methods to study DNA repair pathway choices

To quantify the DNA double strand repair outcomes, different methods have been developed including PCR amplification, followed by either direct sequencing of the modified region or, if the sequence changes containing restriction enzyme recognition sites, restriction-fragment length

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polymorphism (RFLP) analysis (Ran, Hsu et al. 2013).SURVEYOR nuclease assay is also a popular method to quantify the NHEJ events, which is based on the SURVEYOR nuclease cleavage of the reannealed heteroduplexes resulted from indels of NHEJ DNA repair (Ran, Hsu et al. 2013). In addition, targeted genome modifications can be detected by deep sequencing or other next generation sequencing techniques (Ran, Hsu et al. 2013). Several reporter systems have also been developed for the direct measurement of the repair events without sequencing, including DR-GFP reporter, BFP reporter, and traffic light reporter system (Vriend, Jasin et al. 2014, Richardson, Ray et al. 2016).

In this work, the traffic light reporter (TLR) system has been used to monitor DNA repair activities, which allows rapid observation of repair pathway choices (HR or NHEJ) in cells based on fluorescence microscopy and flow-cytometric analysis (FACS).The TLR system consists of a non-functional green fluorescent protein (GFP), followed by a self-cleaving T2A peptide and a second red fluorescent protein (mCherry) in a reading frame shifted by 2 bp (Certo, Ryu et al. 2011). The GFP cDNA sequence contains an insertion comprising an I-SceI site and a stop codon, which disrupts the normal genetic function. Upon repair of the DSB induced around the stop codon, different fluorescent signals will appear depending on whether NHEJ or, in the presence of a DNA template, HDR takes place. Mutagenic NHEJ causes insertions and deletions, thus shifting the downstream mCherry sequence in frame resulting in a red fluorescent (mCherry) signal, whereas homology-directed repair restores the GFP with the help of the DNA donor template, resulting in a green fluorescent signal (Certo, Ryu et al. 2011) (Fig. 12).

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Fig. 12: The traffic light reporter system. The TLR system contains a non-functional GFP sequence, and a

second mCherry sequence in a reading frame shifted by 2 bp. In the GFP cDNA sequence, an I-SceI site and a stop codon was inserted resulted in disrupting normal genetic function. Once a DSB is induced around the stop codon, it can be repaired through either NHEJ (mCherry) or in the presence of a DNA donor template, HDR (GFP) (Certo, Ryu et al. 2011).

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1.4 Aim

The aim of this work is to optimize DNA double strand break repair induced by specific CRISPR-Cas9 nucleases using the Traffic Light Reporter (TLR) system for the development of homologous recombination based gene therapy. To improve the HDR rates, double-stranded DNA plasmid, linearized plasmid sequence, as well as PCR product will be used as donor template with varied homology sequence overlap on the 5’ and 3’ side of the mutation site. Furthermore, repair pathway components will be regulated through chemical or genetic manipulation to bias repair outcomes toward HDR.

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2. Material and methods

2.1 Material

2.1.1 Chemicals and reagents

All chemicals and reagents listed in Table 1 were of high purity grade.

Table 1: Chemicals and reagents

Name Manufacturer

Accutase PAN

Agarose seakem LE Biozym

Ampicillin Sigma-Aldrich

Boric acid Roth

Dimethylsulfoxid (DMSO) Sigma-Aldrich

Disodium phosphate (Na2HPO4) Merck

Dulbecco's Modified Eagle Medium (DMEM) high glucose PAN

DNA loading buffer Fermentas

Ethanol(C2H6O) Roth

Ethidium bromide (EtBr) Roth

Ethylenediaminetetraacetic acid (EDTA) Roth

Fetal bovine serum (FBS) PAN

GelRed Genaxxon

Geneticin Life Technologies

Glycerin(C3H8O3) Merck

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Isopropanol (C3H8O) Roth

Kanamycin Sigma-Aldrich

L-Glutamine PAN

Lipofectamine® LTX and PLUSTM Reagents Invitrogen

Luria broth base Invitrogen

Monopotassium phosphate (KH2PO4) Merck

Penicillin-Streptomycin Biochrom

Potassium chloride (KCl) Roth

Select agar Invitrogen

Sodium chloride (NaCl) Merck

Sodium hydroxide (NaOH) Roth

Tris(hydroxymethyl) aminomethane Roth

2.1.2 Buffers

All buffers were prepared with distilled water from the Sartorius water purification system.

Table 2: Buffers

Name Components

10x TBE 890 mMTris

890 mM Boric acid 20 mM EDTA

Dissolved in deionized water

10x PBS 1.37 M NaCl

27 mMKCl

100 mM Na2HPO4

18 mM KH2PO4

Dissolved in deionized water, PH adjusted to 7.2 with HCl Sterilized and autoclaved

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2.1.3 Media

DMEM (+++) was prepared for eukaryotic cells and LB medium was prepared for prokaryotic cells. Table 3: Media Name Components DMEM (+++) DMEM 10% FBS 2 mM L-glutamine 50 U/mL Penicillin-Streptomycin

LB 25 g Luria broth base

Dissolved in 1 L deionized water Autoclaved

2.1.4 Plasmids

Table 4: Plasmids and expression vectors

Name Provider pcDNA3.1 (+) Invitrogen pcDNA3.1 (-) Invitrogen pcDNA5/FRT Invitrogen px459 (#48139) Addgene hCas9 (#41815) Addgene

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2.1.5 Oligonucleotides

Oligonucleotides listed in Table 5 were obtained from Metabion (Planegg-steinkirchen, Germany) and used for PCR or Sanger sequencing.

Table 5: Oligonucleotides Nr. Name Sequence (5’→3’) 2493 TLR-2a CGCAAATGGGCGGTAGGCGTGTACG 2494 TLR-2b TAGAAGGCACAGTCGAGGCTGATCA 2495 TLR-S1 GAGGGTGGGCCAGGGCACGGGCAGC 2496 TLR-S2 GACCATGGGCTGGGAGGCCTCCTCC 2497 TLR-S3 GCAACTACAAGACCCGCGCCGAGGT 2498 TLR-S4 CTTGTAGGTGGTCTTGACCTCAGCG

2887 eGFP inf1a GAGGGCGAGGGCGATGCCACCTACGGCAAGCTGAC

2888 eGFP inf1b GCATCGCCCTCGCCCTCGCCGGACACGCTGAACTT

2907 delT inf1a TACCCTGTTATCCCTACGCAAAAAGAGCTCACCTACGGC

2908 delT inf1b GCCGTAGGTGAGCTCTTTTTGCGTAGGGATAACAGGGTA

2909 delTAinf1a TACCCTGTTATCCCTACGCAAAAGAGCTCACCTACGGC 2910 delTAinf1b GCCGTAGGTGAGCTCTTTTGCGTAGGGATAACAGGGTA 2924 dGFP 1a ATGCAGAGGCTCCGGTGCCCGTCAG 2925 dGFP 1b AGGGCCCGGATTCTCCTCCACGTCA 2926 dGFP inf1a GAGAATCCGGGCCCTGTGAGCAAGGGCGAGGAGCTGTTCA 2927 dGFP inf1b CCGGAGCCTCTGCATTTACTTGTACAGCTCGTCCATGCCG 2928 dGFP S1 AGAACACCCCCATCGGCGACGGCCC 2933 MY19FS GAGAATCCGGGCCCTAGCGAGCTGATTAAGGAGAACATGC 2934 MY20FS GAGCCTCTGCATTCAATTAAGCTTGTGCCCCAGTTTGCTA

2964 Cas Seq1 GTTTTAAAATGGACTATCATATGC

2965 Cas Seq2 CGCCATCCTGCTGAGCGACATCCTG

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2. Material and methods 34 2967 TLR RS55f ACTTGGCAGTACATCTACGTATTAG 2968 TLR RS55r TTGTCGGGCAGCAGCACGGGGCCG 2969 TLR RS37f AATGGGCGGTAGGCGTGTACGGTGG 2970 TLR RS37r CCCTTGCTCACAGGGCCCGGATTC 2971 TLR RS73f CGTCAATAATGACGTATGTTCCC 2972 TLR RS73r CTTGAAGTCGATGCCCTTCAGCTCG 3464 px459 bf GAATTCTAACTAGAGCTCGCTGATC 3465 px459 br TGGGCCAGGATTCTCCTCGACG 3466 mRFP f GGAGAATCCTGGCCCAGCCTCCTCCGAGGACGTCATCAAG 3467 mRFP r CTCTAGTTAGAATTCTTAGGCGCCGGTGGAGTGGCGGCCC 3242 px459T3f GGTGAGCTCTTATTTGCGTAGTTTTAGAGCTAGAAATAGCAA G 3243 px459T3r TACGCAAATAAGAGCTCACCGGTGTTTCGTCCTTTCCACAAG 3244 px459T4f GGGATAACAGGGTAATGTCGGTTTTAGAGCTAGAAATAGCA AG 3245 px459T4r CGACATTACCCTGTTATCCCGGTGTTTCGTCCTTTCCACAAG 3246 px459T5f GGCCACAAGTTCAGCGTGTCGTTTTAGAGCTAGAAATAGCA AG 3247 px459T5r GACACGCTGAACTTGTGGCCGGTGTTTCGTCCTTTCCACAAG 3305 px459T6f GCCGTCCAGCTCGACCAGGAGTTTTAGAGCTAGAAATAGCA AG 3306 px459T6r TCCTGGTCGAGCTGGACGGCGGTGTTTCGTCCTTTCCACAAG 3307 px459T7f GAGCGCCACCATGGTGAGCAGTTTTAGAGCTAGAAATAGCA AG 3308 px459T7r TGCTCACCATGGTGGCGCTCGGTGTTTCGTCCTTTCCACAAG 3309 px459T8f GGCCGGACACGCTGAACTTGGTTTTAGAGCTAGAAATAGCA AG 3310 px459T8r CAAGTTCAGCGTGTCCGGCCGGTGTTTCGTCCTTTCCACAAG 3238 gRNAT3f TTTCTTGGCTTTATATATCTTGTGGAAAGGACGAAACACCGG

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2. Material and methods 35 TGAGCTCTTATTTGCGTA 3239 gRNAT3r GACTAGCCTTATTTTAACTTGCTATTTCTAGCTCTAAAACTAC GCAAATAAGAGCTCACC 3067 gRNAT4f TTTCTTGGCTTTATATATCTTGTGGAAAGGACGAAACACCGG GATAACAGGGTAATGTCG 3068 gRNAT4r GACTAGCCTTATTTTAACTTGCTATTTCTAGCTCTAAAACCG ACATTACCCTGTTATCCC 3240 gRNAT5f TTTCTTGGCTTTATATATCTTGTGGAAAGGACGAAACACCGG CCACAAGTTCAGCGTGTC 3241 gRNAT5r GACTAGCCTTATTTTAACTTGCTATTTCTAGCTCTAAAACGA CACGCTGAACTTGTGGCC 3252 gRNAT6f TTTCTTGGCTTTATATATCTTGTGGAAAGGACGAAACACCGC CGTCCAGCTCGACCAGGA 3253 gRNAT6r GACTAGCCTTATTTTAACTTGCTATTTCTAGCTCTAAAACTCC TGGTCGAGCTGGACGGC 3254 gRNAT7f TTTCTTGGCTTTATATATCTTGTGGAAAGGACGAAACACCGA GCGCCACCATGGTGAGCA 3255 gRNAT7r GACTAGCCTTATTTTAACTTGCTATTTCTAGCTCTAAAACTGC TCACCATGGTGGCGCTC 3256 gRNAT8f TTTCTTGGCTTTATATATCTTGTGGAAAGGACGAAACACCGG CCGGACACGCTGAACTTG 3257 gRNAT8r GACTAGCCTTATTTTAACTTGCTATTTCTAGCTCTAAAACCA AGTTCAGCGTGTCCGGCC 3298 gesamtTLRf GTTGACATTGATTATTGACTAGT 3299 gesamtTLRr CAGCTGGTTCTTTCCGCCTCAGAAG 3329 RS100Af ACTGGACGGCGACGTAAACGGCCAC 3330 RS100Ar ACGGTGGTGCAGATGAACTTCAGGG 3417 CasSeqf GAGGGCCTATTTCCCATGATTCC 3418 CasSeqr GAGAGTGAAGCAGAACGTGGGGC

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2. Material and methods 36 3419 GFPseqf CTCCGCCCCATTGACGCAAATGGG 3420 GFPseqr GGATGTTGCCGTCCTCCTTGAAGTC 3191 gRNAseqf GATGCATGCTCGAGCGGCCGCCAG 3192 gRNAseqr GAGTTAGCTCACTCATTAGGCACC

2.1.6 Enzymes

All the restriction enzymes listed in Table 6 were used with the recommended buffers according to the manufacturer’s manual.

Table 6: Restriction enzymes

BamHI New England Biolabs (NEB)

BbsI New England Biolabs (NEB)

DraIII New England Biolabs (NEB)

EcoRI New England Biolabs (NEB)

EcoRV New England Biolabs (NEB)

HindIII New England Biolabs (NEB)

NcoI New England Biolabs (NEB)

NdeI New England Biolabs (NEB)

NheI New England Biolabs (NEB)

NotI New England Biolabs (NEB)

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The polymerases listed in Table 7 were used for PCR reactions.

Table 7: Polymerases

PrimeSTAR® HS DNA Polymerase Takara

Phusion® High-Fidelity DNA Polymerase New England Biolabs (NEB)

2.1.7 Markers

Markers listed in Table 8 were used for DNA gel electrophoresis.

Table 8: Markers

GeneRuler 100bp Plus DNA Ladder Fermentas

GeneRuler 1kb DNA Ladder Fermentas

2.1.8 Kits

All the kits listed in Table 9 were used for DNA purification.

Table 9: DNA purification kits

NucleoSpin® Plasmid Macherey-Nagel

QIAfilter Plasmid Midi Kit Qiagen

Qiagen Plasmid Maxi Kit Qiagen

NucleoSpin® Gel and PCR Clean-up Macherey-Nagel PureLink® Genomic DNA Mini Kit Invitrogen

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All the kits listed in Table 10 were used for DNA cloning.

Table 10: DNA cloning kits

In-Fusion® HD Cloning Kit Clontech

TOPO® TA Cloning® Kit Invitrogen

Zero Blunt® TOPO® PCR Cloning Kit Invitrogen

2.1.9 Bacterial strains

Table 11: Bacterial strains

One Shot® TOP10 Chemically Competent E. coli Invitrogen One Shot® TOP10 Electrocomp™ E. coli Invitrogen

Stellar™ Competent Cells Clontech

2.1.10 Devices

Table 12: Devices Name Manufacturer Incubator Binder Autoclave DX-65 Systec GmbH Microscopy VWR

Laminar air flow Invitrogen

BioDocAnalyze Biometra

BioPhotometer Eppendorf

BD Canto II BD

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Centrifuge 1-15 PK Sigma

Centrifuge AK15 Sigma

CO2 incubator Binder

Electrophoresis power supply Biometra

Electrophoresis chambers Biometra

Fluorescence Microscopy Keyence

Gel chamber Whatman Biometra

Thermoblock Biometra

Vortex VWR

Ice machine Scotsman

Magnetic stirrer IKA

PCR-Cycler T Professional Basic Gradient Biometra

PH meter Mettler-Toledo GmbH

Shaker Certomat H Sartorius

Thermoblock Biometra

Scale Ohaus

Water bath TW12 Julabo

2.2 Methods

2.2.1 PCR

PCR (Polymerase chain reaction) is a biological molecular method to amplify a particular DNA sequence through the thermal cycling. The basic components of the PCR include:

1. DNA template, containing the target DNA region to be amplified; 2. two primers, complementary to the 3’ ends of the DNA target region;

Optimizing DNA double strand break repair for homologous recombination based gene therapy